Analysis of Intraflagellar Transport in C. elegans Sensory Cilia

CHAPTER 13

Analysis of Intraflagellar Transport in C. elegans Sensory Cilia Limin Hao*, Seyda Acar*, James Evans*, Guangshuo Ou† and Jonathan M. Scholey* *

Molecular and Cell Biology, University of California, Davis, California 95616

†

Department of Cellular and Molecular Pharmacology, University of California, San Francisco, California 94107

Abstract I. Introduction A. Intraflagellar Transport (IFT) and Cilium Biogenesis on C. elegans Chemosensory Neurons B. Genetic Screens of C. elegans IFT Mutants C. Examination of Ciliary Structure D. In Vivo IFT Motility Assay E. Biochemical Analysis of IFT Motors and Particles II. Rationale A. Genetic Screens of C. elegans IFT Mutants B. Examination of Ciliary Structure C. In Vivo IFT Motility Assay D. Purification of Heterologously Expressed Motor Proteins and In Vitro Motility Assay III. Methods A. Maintenance of WT and dyf Mutant Worms B. Genetic Screens of dyf Mutants and Dyf Assay C. Examination of Ciliary Structure D. Microscopy for In Vivo IFT Motility Assay and Cilia Imaging and Cilia Length Measurement E. Expression and Purification of Kinesin-II and Gliding Motility Assay IV. Materials A. Maintenance of WT and dyf Mutant Worms B. Genetic Screen of dyf Mutants and Dyf Assay C. Examination of Ciliary Structure D. In Vivo IFT Motility Assay and Cilia Imaging E. Purification of Kinesin-II and Gliding Motility Assay METHODS IN CELL BIOLOGY, VOL. 93 Copyright
2009 Elsevier Inc. All rights reserved.

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V. Discussion A. Genetic Screen of IFT Mutants B. Examination of Ciliary Structure C. In Vivo IFT Motility Assay D. In Vitro Gliding Motility Assay VI. Summary Acknowledgments References

Abstract Cilia are assembled and maintained by intraflagellar transport (IFT), the motordependent, bidirectional movement of multiprotein complexes, called IFT particles, along the axoneme. The sensory cilia of Caenorhabditis elegans represent very useful objects for studying IFT because of the availability of in vivo time-lapse fluorescence microscopy assays of IFT and multiple ciliary mutants. In this system there are 60 sensory neurons, each having sensory cilia on the endings of their dendrites, and most components of the IFT machinery operating in these structures have been identified using forward and reverse genetic approaches. By analyzing the rate of IFT along cilia within living wild-type and mutant animals, two anterograde and one retrograde IFT motors were identified, the functional coordination of the two anterograde kinesin-2 motors was established and the transport properties of all the known IFT particle components have been characterized. The anterograde kinesin motors have been heterologously expressed and purified, and their biochemical properties have been characterized using MT gliding and single molecule motility assays. In this chapter, we summarize how the tools of genetics, cell biology, electron microscopy, and biochemistry are being used to dissect the composition and mechanism of action of IFT motors and IFT particles in C. elegans.

I. Introduction A. Intraflagellar Transport (IFT) and Cilium Biogenesis on C. elegans Chemosensory Neurons Cilia (also termed flagella) consist of microtubule (MT)-based axonemes surrounded by a specialized membrane that project from the surface of most eukaryotic cells, where they play vital roles in cell motility, sensory perception, and signaling. The assembly, maintenance, and functions of cilia depend on IFT, a process during which MT-based kinesin and dynein motors drive the bidirectional movement of multiprotein complexes called IFT particles, together with associated cargo molecules, between the base and the tip of the axoneme (Rosenbaum and Witman, 2002; Scholey, 2003). IFT was originally observed in Chlamydomonas reinhardtii (Kozminski et al., 1993) and

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subsequently the IFT particles were isolated and found to consist of two subparticles, IFT subcomplex A (IFT-A) and IFT subcomplex B (IFT-B). IFT-A was resolved into 6 polypeptides and IFT-B into 11 polypeptides (Cole et al., 1998; Piperno and Mead, 1997). The motors that drive anterograde IFT belong to the kinesin-2 family, the founding member of which was identified in sea urchin eggs and embryos as a heterotrimeric, plus-end directed MT-based motor that is required for ciliogenesis (Cole et al., 1993; Morris and Scholey, 1997). It is now known that heterotrimeric kinesin-2 motors (e.g., kinesin-II and the KIF3 complex) drive IFT in most cilia, but its activity can be augmented by homodimeric kinesin-2 motors (e.g., OSM-3 and Kif17) in cilia that contain singlet axoneme extensions (Cole et al., 1998; Hirokawa et al., 2006; Insinna and Besharse, 2008; Scholey, 2008; Snow et al., 2004). Retrograde IFT is driven by the cytoplasmic dynein class 1b, also known as IFT dynein (Schafer et al., 2003; Scholey, 2008; Signor et al., 1999a). The Caenorhabditis elegans genome encodes orthologs of most of the IFT particle and motor polypeptide components (Hao and Scholey, 2009). C. elegans has 60 ciliated sensory neurons whose ciliated dendritic endings are arranged into distinct sensory organs, two amphids in the head and two phasmids in the tail, which are amenable to observations of IFT and sensory ciliogenesis. The sensory cilia on these neurons are approximately 7-µm long and are differentiated longitudinally into three domains: the transition zone, the middle segment, and the distal segment, within which the axonemes display distinct morphologies. The transition zone harbors the basal body which consists of nine triplet microtubules. The middle segment consists of nine doublet microtubules and the distal segment consists of nine singlet microtubules (Evans et al., 2006; Perkins et al., 1986). The techniques available in C. elegans, most notably forward and reverse genetics, genetic manipulations that permit fluorescent protein labeling and worm transformation, and in vivo IFT motility assays, are allowing researchers to study the composition and mechanism of action of IFT motors, IFT particles, and associated IFT components in considerable detail (Fig. 1).

B. Genetic Screens of C. elegans IFT Mutants Following Sydney Brenner’s pioneering work, which was aimed at understanding the genetic control of animal development and behavior using C. elegans as a model system, a variety of behavioral mutants were isolated by ethyl methane sulfonate (EMS)-mediated mutagenesis and were found to affect behaviors such as chemotaxis (che) (Lewis and Hodgkin, 1977), osmotic avoidance (osm) (Dusenbery et al., 1975), and dauer larva formation (daf) (Riddle, 1977). Electron microscopic reconstruction of the sensory neurons in the head of the animal revealed that some of these mutants exhibited structural defects of sensory cilia located on the dendritic endings of sensory neurons (Perkins et al., 1986). Dye-filling assays demonstrated that the mutants displaying defective cilia structure could not absorb fluorescent dye from the surrounding environment into their amphids and phasmids, whereas the

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(A)

(B)

Fig. 1 The IFT machinery of C. elegans sensory cilia. (A) All the IFT particle components are arranged into three complexes: IFT-A, IFT-B, and BBSome complex. The components in the circles in IFT-A and IFTB correspond to those originally isolated from C. reinhardtii. Other subunits were observed to move together with these subunits undergoing IFT. The arrangement of the BBS components in the BBSome complex is based on Nachury et al. (2007). (B) The subunit compositions of the two anterograde IFT motors, the heterotrimeric kinesin-II and homodimeric OSM-3, and the retrograde motor IFT dynein from C. elegans. IFT dynein probably contains additional light and intermediate subunits.

wild type could do so (Perkins et al., 1986). In an independent dyf (dye-filling defective) mutant screen, 13 dyf mutants representing new genetic loci were isolated, and the molecular lesions responsible for the mutant phenotype associated with most of them have subsequently been elucidated (Table I) (Starich et al., 1995). During the same year that IFT was discovered in C. reinhardtii, and the IFT motor, kinesin-2 was purified from sea urchin embryos, osm-3, was cloned as a ciliary mutant and was deduced to encode a kinesin-like protein (Shakir et al., 1993). Subsequently, the osm-6 mutant was cloned (Collet et al., 1998) and found to encode an IFT52 homologue, a subunit of the IFT-B subcomplex (Cole et al., 1998; Piperno and Mead, 1997). Thus, early C. elegans genetic analysis of animal behavior provided a foundation for the subsequent analysis of ciliary biogenesis and IFT using this organism. To complement forward genetics, transposon insertion mutagenesis, chemical and UV-TMP (trimethylpsoralen) mutagenesis, and PCR-based protocols have been applied to the isolation of deletion alleles of any given genes in C. elegans (Jansen et al., 1997; Liu et al., 1999; Zwaal et al., 1993). This reverse genetic approach is particularly critical for studying sensory ciliogenesis and IFT because the RNAi technique which is so effective in other systems has not been shown to robustly knockdown genes in ciliated sensory neurons, even in sensitized mutant backgrounds. Moreover, the C. elegans gene knockout consortium facility uses UV-TMP mutagenesis and PCR screens to generate mutations in any gene of interest, in response to researchers’ requests (http://celeganskoconsortium.omrf.org/ or http://www.shigen.nig. ac.jp/c.elegans/).

Name che-2 che-3 che-11 che-13 daf-10 daf-19 dyf-1 dyf-2 dyf-3 dyf-4 dyf-5 dyf-6 dyf-7 dyf-9 dyf-10 dyf-11 dyf-12 dyf-13 osm-1 osm-3 osm-5 osm-6 osm-12

Other name

osm-2, che-8, avr-1, caf-2 che-9 osm-4 daf-24

dyf-8

caf-1, klp-2

bbs-7

Gene model/Genetic position (cM)

che

daf

dyf

osm

Function

References

F38G1.1 F18C12.1 C27A7.4 F59C6.7 F23B2.4 F33H1.1a F54C1.5a ZK520.1 C04C3.5a V:4.31 ± 0.29 M04C9.5 F46F6.4 C43C3.3 V:24.22 ± 0.35 I:1.53 ± 0.040 C02H7.1 X:2.18 ± 0.07 C27H5.7a T27B1.1 M02B7.3a Y41G9A.1 R31.3

þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ

þ þ þ þ þ þ / / / / / / þ þ / / þ þ þ þ þ þ

þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ þ

þ þ þ þ þ þ þ / / / / / þ / / / / / þ þ þ þ

IFT-particle B IFT-dynein heavy chain IFT-particle A IFT-particle B IFT-particle A RFX family transcription factor OSM-3-kinesin activator IFT-particle IFT-particle B associated

Fujiwara et al. (1999) Wicks et al. (2000) Qin et al. (2001) Haycraft et al. (2003) Bell et al. (2006) Swoboda et al. (2000) Ou et al. (2005b) Efimenko et al. (2006) Murayama et al. (2005)

MAP kinase IFT-particle Cell-surface ligand

Burghoorn et al. (2007) Bell et al. (2006) Heiman and Shaham (2009)

IFT-particle B associated

Bacaj et al. (2008)

Y75B8A.12

þ

/

þ

þ

Ciliary distal segment assembly IFT-particle B IFT-kinesin IFT-particle B IFT-particle B Part of ciliary distal segment assembly

Blacque et al. (2005) Bell et al. (2006) Shakir et al. (1993) Haycraft et al. (2001) Collet et al. (1998) Blacque et al. (2004)

13. Analysis of Intraflagellar Transport in C. elegans Sensory Cilia

Table I C. elegans Ciliary Mutants

Keys: che, chemotaxis assay; daf, daufer formation assay; dyf, dye-filling assay; osm, osmotic avoidance assay.

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C. Examination of Ciliary Structure The cilia present on the dendrites of C. elegans chemosensory neurons are fasciculated and localized along the channel formed by two glia-like cells, namely the sheath cell and socket cell. These channel cilia are in direct contact with the environment, which facilitates the sensing of environmental cues (Lewis and Hodgkin, 1977; Ward et al., 1975). When the worms are immersed in a lipophilic fluorescent dye, for example, FITC or DiI, a subset of the channel cilia uptake the dye, producing fluorescently stained sensory neurons. Consequently, sensory neurons that fail to uptake the fluorescent dye have perturbations of their ciliary structure, often due to defects in cilium assembly or maintenance. The ultrastructural analysis of such dyf mutants by transmission electron microscopy (TEM) has indeed revealed all kinds of ciliary structural defects (Perkins et al., 1986). The subsequent molecular analysis of these mutants demonstrated that many of them are caused by mutations in IFT components. Serial section TEM can be employed to confirm that morphology defects of IFT motor mutants observed using light microscopy reflect a change in the ultrastructure of the amphid channel cilia (Chalfie and Thomson, 1982; Evans et al., 2006; Perkins et al., 1986). While the acquired images only represent a snapshot of the cross-sectional view every 50–150 nm along their length, a model of the path of each cilium can be generated and used to help understand the role of specific gene products in IFT and cilium biogenesis. Aligning images of serial sections corresponding to the first 10 µm from the tip of the head of the worm allows the tracking and modeling of individual cilia from their neuronal transition zone to their exposed endings at the amphid pore (Fig. 2A). Additionally, if the micrographs are recorded on film or CCD at a sufficient resolution (~2 nm/pixel) then the architecture of each axoneme can also be determined with the MTs arranged either as singlets, doublets, or triplets (Fig. 2A). Reconstruction of serial transmission sections of the worm’s head allows researchers to observe the morphology and the length of the cilia, the architecture of the axoneme, and other details. In most cases though, mutations in IFT genes cause significant shortening of cilia which can be seen by introducing a fluorescently labeled cilia marker into the mutant allowing observations of cilium length under fluorescence microscopy (Fig. 2B). By correlating results from light and TEM, it has been shown that two kinesin-2 motors, homodimeric OSM-3 kinesin and heterotrimeric kinesin II, function in a partially redundant manner to build full-length amphid channel cilia but are completely redundant for building full-length amphid wing (AWC) cilia. This difference apparently reflects cilia-specific differences in OSM-3 activity, which serves to extend distal singlets in channel cilia but not in AWC cilia which lack such singlets and respond to different stimuli (Chalfie and Thomson, 1982; Evans et al., 2006; Perkins et al., 1986). D. In Vivo IFT Motility Assay Using time-lapse fluorescence microscopy, the movement of specifically labeled IFT proteins along the sensory cilium can be readily visualized in C. elegans. First,

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green fluorescent protein (GFP)-labeled kinesin-II (tagged on the KAP-1 subunit) and OSM-6, an IFT-B component, were observed to move along the cilium (Orozco et al., 1999). The same technique was subsequently applied to other IFT particle components, leading to the identification and characterization of a number of IFT-A and IFTB components such as OSM-1, OSM-5, CHE-2, CHE-11, DAF-10 (Qin et al., 2001). Later, several more IFT particle components, including IFTA-1 (Blacque et al., 2006), DYF-2 (Efimenko et al., 2006), and CHE-13 (Haycraft et al., 2003) were identified and characterized (Fig. 1A). Because the fluorescent signal emitted by GFP-tagged IFT components accumulated in the ciliary tips of che-3 and xbx-1 mutants, these latter two genes were identified as subunits of dynein 1b, the retrograde IFT motor (Schafer et al., 2003; Signor et al., 1999a) (Fig. 1B). With the introduction of spinning disc confocal microscopy and kymography, the IFT assay became a routine method for studying the mechanism of IFT in this system (Snow et al., 2004) (Fig. 3). In addition to kinesin-II, OSM-3 was found to contribute to transport along the cilium by coordinating with kinesin-II to transport IFT particles along the middle segment and to move along and build the distal segment on its own (Evans et al., 2006; Snow et al., 2004). Kinesin-II is a slow motor, moving at 0.5 µm/s in the absence of OSM-3, whereas OSM-3 is a fast motor, moving at 1.3 µm/s in the absence of kinesin-II (Snow et al., 2004). In wild-type animals, kinesin-II and OSM-3 cooperate to move the same IFT particles along the middle segment at a velocity intermediate between those of the individual motors (0.7 µm/s) whereas OSM-3 moves along the distal singlets on its own at its characteristic fast velocity (1.3 µm/s). By analyzing the rates of IFT in bbs (Bardet–Biedl syndrome) mutants, it was found that kinesin-II moves at its characteristic slow rate (0.5 µm/s) and OSM-3 moves at its characteristic fast rate (1.3 µm/s) along the axoneme middle segment, suggesting that BBS complex is responsible for the coordination of the two anterograde IFT motors (Ou et al., 2005b). Several GFP-tagged IFT-A components move at the slow kinesin-II rate (0.5 µm/s) whereas IFT-B components move at the fast OSM-3 rate (1.3 µm/s), indicating that kinesin-II carries the IFT-A subcomplex and OSM-3 carries the IFT-B subcomplex (Ou et al., 2005b; Pan et al., 2006) (Table II). In combination with the analysis of dyf mutants, four novel IFT-B components, DYF-1 (Ou et al., 2005b), DYF-3 (Murayama et al., 2005; Ou et al., 2005a), DYF-11 (Omori et al., 2008), and DYF-13 (Blacque et al., 2005) were identified and characterized; these components had been missed using the powerful approach of C. reinhardtii biochemistry, underscoring the value of using complementary techniques in multiple systems (Cole et al., 1998; Piperno and Mead, 1997). The putative cargo of the IFT machinery was also studied assuming that they may be cotransported with the IFT machinery, for example, OSM-9 and OCR-2 (Qin et al., 2005). This powerful approach also allowed investigators to identify dyf-5, a map kinase, as a modulator of the velocity of OSM-3 (Burghoorn et al., 2007). E. Biochemical Analysis of IFT Motors and Particles Taking advantage of the ease of cell culture and deflagellation, IFT particles have been isolated from C. reinhardtii flagella and their components have been nicely

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(A)

Distal

Middle

Transition

klp-11

osm3/ klp-11

3-D Model

Wild Type

kap-1/ osm-3

kap-1

klp-11

klp-11/ osm-3

kap-1/ klp-11

osm-3

che-3

(B) WT

Fig. 2 (Continued)

osm-3

che-3

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characterized. Antibodies raised against separated IFT components were used for further identification of flagellar mutants and for further investigation of the localization and mechanism of action of IFT components. In addition, by changing the ionic strength, it was determined that isolated IFT particles could dissociate into two subcomplexes, IFT-A and IFT-B, of which IFT-B was found to be formed from a core particle to which a few additional loosely associated polypeptides are bound (Fig. 1A). Recently, the “BBSome,” another subcomplex associated with the IFT machinery, has also been isolated using tandem affinity purification from cultured mammalian cells. This body of work provides useful insights into the native form of the multi-protein complex involved in IFT. However, although most, if not all, the components of the IFT particle have been identified, our understanding of how these IFT components interact with each other and how they interact with motors and cargos is relatively poor (Hao and Scholey, 2009). Unfortunately, C. elegans is not amenable for the biochemical isolation of endogenous IFT particles since it contains few ciliated cells (60 out of ~1000 cells) and the cilia themselves are tiny. Therefore, it is necessary to heterogeneously express IFT proteins in order to investigate the interactions and biochemical properties of these recombinant proteins in vitro. OSM-3, one of the two anterograde IFT motors, has been overexpressed in an Eschericia coli system where both a GFP and a 6 His Tag have been used to facilitate the protein’s purification. The purified recombinant protein was used for sophisticated in vitro gliding motility assays, ATPase assays, single molecule fluorescence, and optical trap experiments (Imanishi et al., 2006; Pan et al., 2006). Heterotrimeric kinesin-2 was initially purified from sea urchin by using pan-kinesin peptide antibodies raised against hyperconserved sequences within the kinesin superfamily motor domain (Cole et al., 1992; Cole et al., 1993). Later purification trials of the orthologous kinesin-II complex from C. elegans resulted only in its partial purification due to the low levels of the native holoenzyme, although it was possible to use coimmunoprecipitations and fractionation procedures to determine its heterotrimeric state, and to reveal that OSM-3 formed a distinct, homodimeric complex (Signor et al., 1999b). Recently we have described the heterologous expression of genes encoding klp-11, klp-20, and kap-1, the three subunits of kinesin-II simultaneously in the baculovirus expression system using Sf9 cell cultures (Pan et al., 2006). The production of heterologously expressed kinesin-II in bulk amounts compared to native kinesin-II has allowed us to isolate the protein in high-purity relatively easily. The

Fig. 2 Examination of ciliary structure. (A) TEM analysis of amphid channel cilia in IFT motor mutants. Representative micrographs of distal and middle segments and transition zones and a side view of the corresponding 3D model that was reconstructed from serial sections from each strain. Three-dimensional reconstructions show trajectories of channel cilia obtained from serial sections taken from the distal tip at the amphid pore down through the basal transition zones to the dendrites (total length ~10 µm). Bars, 250 nm. (B) Amphid channel cilia morphology analyzed by fluorescence microscopy. Amphid channel cilia visualized with IFT particle proteins, OSM-5::GFP or OSM-6::GFP. Arrowheads indicate middle (bottom) and distal (top) segment boundaries. (Reprinted from Evans et al., 2006).

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(A)

M1

M2

M3

M4

OSM-6 GFP

D 140

Middle

Distal

Transport Events

120

D

100 80 60 40 20

M1 M2 M3 M4

0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1 1.1 1.2 1.3 1.4 1.5 1.6 1.7 1.8

Velocities (µm/s)

(B) M1

M2

M3

M4

140

OSM-6 GFP in osm-3

Transport Events

120 100 80 60 40 20

M4 M3 M2 M1

0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1 1.1 1.2 1.3 1.4 1.5 1.6 1.7 1.8

Velocities (µm/s)

(C) M1

M2

M3

M4

D

OSM-6 GFP in KAP-1

80

D

Transport Events

70 60 50 40 30 20 10

M4 M3 M1 M2

0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1 1.1 1.2 1.3 1.4 1.5 1.6 1.7 1.8

Velocities (µm/s)

Fig. 3 Anterograde transport of IFT-particles along the middle and distal segments of C. elegans sensory cilia. Motility of IFT particles (OSM-6::GFP) within sensory cilia of wild-type (A), osm-3 mutant (B) and kap-1 mutant (C). Left column show fluorescence micrographs with corresponding cartoon showing the lines used to generate kymographs along 4 middle segments (M1–M4) and the distal segment (D). Kymographs (middle column) show that motility along the distal segments is faster than along middle segments (no distal segment motility is seen in the osm-3 mutant). Right column, histograms showing IFT velocity profiles along middle and distal segments. Horizontal bars = 2.5 µm. Vertical bar = 5 s. (Reprinted from Snow et al., 2004).

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Table II Velocities of the IFT Motors and Particles in the In Vitro Gliding Assay and In Vivo Motility Assay Pure motor (µm/s)

Motor in cilium (µm/s)

IFT particles (µm/s)

kinesin-II

0.4

0.5

0.5

OSM-3

1.1

1.3

1.2

kinesin-II and OSM-3

0.7

0.7

0.7

Assembly of

Middle segment Full-length cilium/ distal segment Middle segment (redundant)

use of the baculovirus expression system for the production of this complex protein is important in that it allows protein expression in a higher eukaryote system which contains the machinery required for post-translational modifications. This in turn can yield recombinant proteins sharing properties with the native form. Kinesin-II and OSM-3 work together to move an IFT particle along the middle segment of the sensory cilium. The coordination of these two motor proteins was reconstituted by assaying the gliding velocities of mixtures of these two proteins in a series of different ratios. This approach also enabled us to closely correlate the rates of motility of the purified motors with the rates of IFT seen in different domains of wildtype and mutant sensory cilia, allowing us to develop a model for the coordination of the two ciliary motor proteins (Table II) (Pan et al., 2006).

II. Rationale A. Genetic Screens of C. elegans IFT Mutants Using forward genetic approaches in C. elegans, gene inactivation is accomplished in two ways. EMS mutagenesis is the most widely used technique to mutate the C. elegans genome (Brenner, 1974). The ability of EMS to alkylate guanine bases results in base mispairing, primarily producing G/C to A/T transitions, which ultimately causes amino acid changes, deletions, or insertions during subsequent DNA repair. Alternatively, Mos1 transposon-mediated mutagenesis has been used to insert Mos1 DNA fragment into the C. elegans genome to generate mutations (Boulin and Bessereau, 2007). EMS- or Mos1-treated animal can be screened for defects in chemotaxis, osmotic avoidance, dauer larva formation, and dye filling. Among these assays, dye-filling assay is very easy to perform and also provides a direct assessment of the presence of ciliary defects (Perkins et al., 1986). Screens for behavioral mutants are also easy to perform and some of the mutants from these screens are indeed required for IFT or ciliogenesis. However, a significant portion of

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these mutants develop normal cilia and they may therefore play other roles, for example, participating in signal transduction in response to environmental stimulation. Therefore, the dye-filling assay needs to be performed to examine their roles in ciliogenesis (Ou et al., 2007). The identification of EMS-induced mutations in the genome is a time-consuming task. The strategy starts with two-point mapping to determine on which chromosome the mutation localizes by genetic crossing with other phenotypic markers. Then three-point mapping is applied to narrow down the locus of the mutation on the chromosome. The recent application of a snip-SNP (restriction site modified single-nucleotide polymorphism) mapping protocol based on single-nucleotide differences between Bristol (the widely used wild type) and Hawaiian strains of C. elegans significantly speeds up the genetic mapping of ciliary mutants (Wicks et al., 2001). When the mutation is mapped to a small region within several hundred kilobases, a pool of cosmids or fosmids carrying wild-type genes covering this region can be individually transformed into mutant animals by germ-line microinjection, in order to test if they can rescue the mutant phenotype. All the genes in the cosmid or fosmid that rescue the mutant phenotype can then be amplified by PCR to examine their ability to rescue the mutant phenotype, or alternatively, the exons of all the genes on the construct can be amplified by PCR and subjected to sequencing in order to identify the mutation. Since genes containing “X-box motifs” in their promoter regions are likely to be ciliary genes, the identification of candidates for further analysis can be facilitated by bioinformatics searches for putative X-box motifs (see the reason in the next paragraph). Compared with EMS mutations, the identification of Mos1 transposon insertion-induced mutation is relatively straightforward. Mos1 insertions represent molecular tags that allow inverse PCR to amplify the flanking sequences of Mos1 in the C. elegans genome. Moreover, with the advancement of “next-generation” sequencing techniques, whole genome sequencing can be used to identify sought-after mutations (Hillier et al., 2008; Sarin et al., 2008). C. elegans reverse genetics is based on the fact that IFT is a process occurring in the cilium which is a compartmentalized space requiring the transport of cytoplasmically synthesized proteins. Thus, genes contributing to IFT are expressed in ciliated sensory neurons and their products are located in cilia. To identify ciliated sensory neuronexpressed genes, a few strategies have been used: (1) All the currently identified IFT genes and many ciliary genes have an X-box motif in their putative promoter region that can be recognized by DAF-19, an RFX family transcription factor, one isoform of which is specifically expressed in ciliated sensory neurons (Swoboda et al., 2000). On the basis of this, genome-wide searches for genes containing X-boxes in their putative promoter regions predict that hundreds of genes could be regulated by DAF-19 in ciliated sensory neurons (Chen et al., 2006; Efimenko et al., 2005). (2) Serial analysis of gene expression (SAGE) data collected from in vitro cultured sensory neurons provide a list of candidate genes that are expressed in C. elegans sensory neurons (Blacque et al., 2005). (3) Overexpression of a GFP-tagged mRNA poly(A)-binding protein (PAB) in ciliated sensory neurons, followed by the use of a GFP antibody to

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isolate the mRNA species that bind to PAB has been used in the identification of the isolated mRNA which provides information about genes that are specifically expressed in ciliated sensory neurons (Kunitomo et al., 2005). (4) The list of the ciliary genes discovered by comparative genomics (Avidor-Reiss et al., 2004; Li et al., 2004) and proteomics (Liu et al., 2007; Mayer et al., 2008, 2009; Ostrowski et al., 2002; Pazour et al., 2005; Song and Sokolov, 2009) in other organisms can be used to identify likely ciliary genes in C. elegans. Once a set of putative candidate genes are available, these genes can be substantiated as IFT components based on the location of their protein product, IFT assays, and mutant analysis. To isolate C elegans deletion mutants in genes of interest, two steps are involved (Jansen et al., 1997). The first step involves random mutagenesis. A large population of worms is irradiated under UV illumination in the presence of trimethylpsoralen which creates small deletions (presumably in all the genes). Half of the mutated worms are frozen and another half of them are used to extract genomic DNA. The second step is to screen for the mutation of interest by PCR. The primers are designed to flank the desired deletion region in the gene of interest. If a deletion occurs in the region, it brings primers closer together and generates a PCR product smaller than wild type. When a deletion is detected in a DNA pool, one can go back to the corresponding frozen worm stock and recover their siblings which bear the same mutation. Recently, oligonucleotide array comparative genomic hybridization has been used as a new tool to identify the deletion. Using exon-tiled oligonucleotide arrays, DNA samples from WT and mutagenized animals are differentially labeled and hybridized to detect singlecopy single-gene deletions (Flibotte et al., 2009; Maydan et al., 2007). In addition, targeting induced local lesion in genomes (Gilchrist et al., 2006) and PCR product sequencing (Cuppen et al., 2007) have also been adopted to identify mutations of interest.

B. Examination of Ciliary Structure The dye-filling defect phenotype of the dyf mutants typically suggests that cilia structure is abnormal (Hedgecock et al., 1985; Perkins et al., 1986). There are two main ways to examine ciliary structure: TEM and fluorescent microscopy. TEM analysis allows direct observations of axoneme architecture and length, as well as cilia membrane structure (Chalfie and Thomson, 1982; Evans et al., 2006; Perkins et al., 1986). Fluorescence microscopy allows analysis of IFT and cilia length based on the distribution of fluorescently labeled markers. The structure of the basal body, ciliary membrane, axoneme, and matrix has been studied using fluorescent labeling of known components of these domains in the cilium. Ciliary membrane components, OSM-9, OCR-2, TUB-1, etc., have been labeled fluorescently and observed in transgenic worms (Mukhopadhyay et al., 2005; Qin et al., 2005). The alpha, beta, and gamma tubulins are components of the axoneme. The tbb-4 gene, coding for a beta tubulin, has been shown to be specifically expressed in the amphids and phasmids, which makes it a perfect marker to analyze ciliary lengths (Jauregui et al., 2008;

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Portman and Emmons, 2004). The tbg-1 gene encodes a gamma tubulin, which makes it a good marker of the basal body (Tabish, 2007). IFT particle components locate to the matrix of the cilia. So far, almost all the IFT components (Fig. 1A and B) have been labeled with GFP, which are perfect for IFT assays and in most cases are also good markers for ciliary length analysis. In addition, GFP diffuses into cilia by itself when it is expressed in a subset of cilia driven by certain sensory neuron-specific promoters, gcy-5, gcy-7, etc. (Yu et al., 1997). However, the diffusible GFP does not fill the entire full-length cilium.

(A)

(B)

(C)

(D)

Fig. 4 (Continued)

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C. In Vivo IFT Motility Assay C. elegans is transparent and each of its cells can be observed directly under a microscope. Fluorescently tagged IFT proteins can be integrated into the IFT machinery and their motility can be recorded. By analyzing the recorded movie by kymography, the moving velocity can be estimated. In order to perform in vivo motility assays of IFT, transgenic worms that carry a fluorescently tagged IFT protein should be available. An appropriate transgene may be constructed by three commonly used methods (Fig. 4). (1) Worm expression vectors that contain GFP can be used by either traditional cloning (Fire et al., 1990) or gateway recombination (Hope et al., 2004). A gene of interest together with its own putative promoter region is amplified by PCR and cloned into the upstream region of GFP in the vector, either by restriction digestion and ligation in conventional genetic engineering or by recombined arms in the gateway recombination system (Fig. 4A and B). (2) Fusion PCR is used to directly fuse a gene of interest together with its putative promoter to the GFP sequence, and the purified PCR product is directly used as a transgene (Hobert, 2002). This method requires two rounds of PCR. First of all, the gene of interest and GFP are amplified, respectively, using one of the primer pairs that are engineered to recognize each other so that the PCR products can overlap within the engineered sequence. Then the two nonoverlapped primers that were used in the first-round PCR are used for the next-round PCR (Fig. 4C). (3) The GFP encoding sequence is recombined into a gene of interest in a fosmid (Dolphin and Hope, 2006). This method requires two rounds of recombination in a genetically modified bacterial system. In the first round, the PCR product of a gene cassette containing rpsL and tetA(C) genes under control of a strong promoter, ompF, is generated with the primers that are engineered to recognize the sequences flanking the insertion point in the gene of interest. It is transformed into bacteria that contain a single copy of the fosmid, recombination is allowed to occur, and successful recombinants are selected for the TetR marker since the two markers confer streptomycin sensitivity (StrS) and tetracycline resistance (TetR) to the bacterial host, respectively. In

Fig. 4 Schematic cartoon of gene cloning strategies used to make GFP-tagged IFT protein constructs. (A) A gene of interest (e.g., osm-3) together with its promoter is amplified by PCR using two primers engineered into restriction sites (RE1 or RE2), respectively, which recognize the same sites in a vector that carries the GFP sequence. (B) The gene of interest (e.g. osm-3) together with its promoter is amplified by PCR with two primers engineered into a recombination site (attL or attR) which recognizes the same site in a vector that carries the GFP sequence. (C) A gene of interest (e.g., osm-3) together with its promoter is amplified by PCR using two primers. The downstream primer is engineered into a sequence that overlaps the 50 end of the GFP sequence. Another GFP sequence, together with the unc-54 30 UTR, is amplified by PCR. Next, the two PCR products are mixed together and the fusion PCR is performed by using two primers: one upstream of osm-3 and another downstream of the unc-54 30 UTR. (D) Recombineering. The PCR product of a double-selection marker [ompF::rpsL::tetA(C)], amplified by two primers engineered in two 50 mer of sequences that are the same as the sequences flanking the insertion point in a gene of interest (e.g., che-3) in a fosmid, is used to perform first-round recombination. The recombinant is selected by TetR because of the insertion of tetA(C) in the che-3 gene. In the second round of recombination, the PCR product of GFP amplified by two primers engineered in two 50-mer sequences that are the same sequences flanking the insertion point is used. The recombinant is selected by StrR because of the removal of rpsL.

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the next round of recombination, the PCR product of GFP containing the flanking recognition sequences at the insertion point is transformed into the recombinant from the first-round screen for streptomycin resistant recombinant (Fig. 4D). To generate transgenic worms, the construct made by either of the above methods together with an injection marker (e.g., a dominant rol-6 mutant allele in pRF4 that confers a rolling phenotype to transgenic worms) are injected into the worm’s gonad. The transgene forms an extrachromosomal array that contains multiple copies of the transgene (Mello et al., 1991). The bombardment of the transgene can also be used to produce transgenic worms carrying a low copy number of the transgene (Praitis et al., 2001). The successfully created transgenic worms are observed under a fluorescence microscope to see if the transgene is expressed, as judged by the existence of a GFP signal. The transgene is also transformed into the corresponding mutant background to see if it rescues the mutant phenotype, and thus to determine if it is functional. Any transgenic animals carrying a functional transgene can be used to create IFT motility movies under a spinning disc confocal microscope. D. Purification of Heterologously Expressed Motor Proteins and In Vitro Motility Assay Testing purified IFT motor proteins for their motility activity and assaying their biochemical properties is obviously important for improving our understanding of the mechanism of IFT. To this end, motility assays of dyneins and kinesins based on time lapse imaging by video-enhanced fluorescence microscopy of rhodamine-labeled MTs are especially convenient. The assay involves attaching motor proteins to a glass slide and assaying motility using buffer solutions containing ATP and taxol stabilized MTs. Using this approach, the C. elegans IFT motors, heterotrimeric kinesin-II, and OSM-3 have been studied (Scholey, 2008). Motility assays were used to determine the motility properties of each protein individually as well as for a series of mixtures of varying molar ratios of the two proteins. In this study, heterotrimeric kinesin-II was expressed using the baculovirus system and purified using affinity chromatography and gel filtration chromatography, whereas the OSM-3 motor was expressed in E. coli and purified in the Vale Lab. The use of mixtures of the two proteins mimicked the in vivo conditions and led to the generation of a model for IFT in C. elegans that explains how kinesin-II and OSM-3 are functionally coordinated to move the same IFT particle along the middle segment of amphid channel cilia (Table II) (Pan et al., 2006).

III. Methods A. Maintenance of WT and dyf Mutant Worms

1. NGM Plate Preparation and Worm Culturing 1. To a 2-l flask with a stir bar, add 975 ml ddH2O, 2.5 g peptone, 3.0 g NaCl. Stir until dissolved, and then add 19 g of bacto-agar and autoclave at 120°C for 15 min.

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2. Allow NGM agar to cool until the flask may be touched by hand (~55°C), and then add the following sterilized solutions: 1 ml cholesterol (5 mg/ml stock in ethanol), 1 ml 1 M MgSO4, 1 ml 1 M CaCl2, 25 ml 1 M Na-Phosphate, pH 6.0. 3. Pour the plates (35 mm10 mm). When the agar hardens, seed the plates with an overnight culture of OP50 bacterial strain. 4. Pick two adult worms and transfer onto the seeded NGM plates and culture the worms at 20°C for 3–4 days until their progeny grow to adult stage. These worms can be used for the IFT assay and ciliary structure analysis.

2. Freezing Worms 1. Culture six plates of worms at 20°C. When the plates are full of worms, wash off the worms with M9 buffer into a 15-ml plastic tube. 2. Centrifuge at 4000 rpm (2200 g) for 1 min to collect the worms. 3. Aspirate the solution till 1.5 ml is left in the tube and then add 1.5-ml freezing buffer and mix. 4. Dispense them into three cryo-vials, 1 ml per vial, and then put the three vials into a Styrofoam rack and incubate in a –80°C freezer. (The Styrofoam rack allows the temperature to fall gradually). 5. When the worms are frozen (1 h or longer), the worms can be stored in the –80°C freezer or in liquid nitrogen. B. Genetic Screens of dyf Mutants and Dyf Assay

1. Isolation of dyf Mutants 1. Culture wild-type (N2 strain) worms at 20°C to get many L4 larval stage worms or young adults. Wash off the worms into a 15-ml tube. Spin down the worms and resuspend in 3 ml M9. 2. Prepare another 15-ml tube with 1 ml M9 and add 20 µl EMS and mix. 3. Combine the worms and EMS. Incubate on a shaker at 20°C for 4 h. 3. Spin down the worms and wash three times with M9 buffer. 4. Transfer the worms onto fresh NGM plates and allow them to recover for 4 h to overnight. 5. Pick four worms (P0) onto each NGM plate and repeat for 60 plates. 6. Culture the worms for 3 days to obtain F1 progeny, and pick four worms onto each plate and repeat for 200 plates or more. 7. Stain the F2 worms with DiI solution. Observe under a fluorescence dissecting microscope to search for worms that are not able to be stained. 8. The putative dyf worms are singled onto fresh plates and bred true.

2. Dyf Assay 1. Culture the worms till the plate is full of worms of different ages. 2. Make the fluorescent dye solution (DiI for red fluorescence or DiO for green fluorescence) by diluting (1:200) the stock solution with M9 buffer.

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3. For a 3.5-cm plate, add 0.5 ml diluted fluorescent dye solution and swirl the plate to make sure all the worms are immersed in the solution, and incubate for 1 h or longer until the plate surface becomes dry. 4. Observe the stained worms directly under a fluorescence dissecting microscope or pick the worms on a 2% agarose pad with a drop of 10 mM NaN3 and observe under a compound microscope with a 60 objective. C. Examination of Ciliary Structure

1. Preservation of Worms 1. Load worms into specimen holder and insert into high-pressure freezer. 2. Using 2000 bar pressure and liquid nitrogen, freeze specimen and transfer under liquid nitrogen into cryo-vial containing substitution cocktail: 1% OsO4, 0.1% uranyl acetate in 100% acetone. 3. Replace the water in the frozen sample with acetone using a freeze substitution device using the following protocol: a. 3 days at –90°C (Leica AFS) b. 12 h at –25°C c. 3–4 h at 4°C d. 1–2 h at room temperature e. Rinse with 99.99% anhydrous acetone 4. Infiltrate and embed worms in Epon-Araldite resin using the gradient approach. a. 3:1 acetone : resin for 2 h b. 2:2 acetone : resin for 4 h c. 1:3 acetone : resin overnight d. 100% resin overnight 5. Polymerize resin at 70°C for 72 h.

2. Sample Preparation for TEM 1. Align the polymerized resin block to obtain the proper orientation of the worm for obtaining cross-sections. 2. Trim the face of the resin block to a trapezoid just larger than the desired specimen area using a handheld razor blade. 3. Load the trimmed resin block into the ultramicrotome cutting arm and slice 50- to 150-nm thick sections using a glass or diamond knife. 4. Collect the floated serial sections on a formvar coated 3 mm copper TEM slot grid. 5. Stain the grid containing serial sections with 2% uranyl acetate dissolved in methanol for 5 min followed by Reynold’s lead citrate for 2 min.

3. Imaging and Modeling 1. Load grid with serial sections into TEM sample holder and insert into microscope. 2. After using standard alignment procedures, acquire transmission electron micrographs of the sample area at a magnification sufficient for resolving desired information.

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3. Due to the resulting gap of 50–150 nm between consecutive sections the images should be manually aligned using the manual image deformation and alignment system contained within IMOD (freely available from http://bio3d.colorado.edu/imod). 4. After the sections are aligned, model the positions of the microtubules, cilia, and other relevant organelles as surface renderings using 3DMOD (within the IMOD package) to depict the length, path, and axoneme architecture of each cilium. D. Microscopy for In Vivo IFT Motility Assay and Cilia Imaging and Cilia Length Measurement

1. Immobilization of Worms 1. Prepare agarose pads: Put a drop of melted 2% agarose in M9 buffer on a slide and cross another slide on it and press gently. When it is used, detach the two slides and the agarose layer is formed on one of them. 2. Get a cover slip and transfer 16 µl of 10 mM levamisole on it and then pick four worms into the levamisole solution and incubate 5 min with a cover preventing evaporation. Adult worms are usually used for the IFT assay because their cilia size is large and easy to be observed. 3. Reverse the agarose pad and put it onto the cover slip with anesthetized worms. 4. Seal the slide assembly with Valap: Dip a cotton stick into the melted Valap and brush along the edges of the cover slip. 5. Look at the worms against the light and label the worms with a marker pen on the slide side of the assembly so as to find the worms easily in the following step under higher magnification (100). 6. Put a drop of oil on the cover slip and reversely put onto the platform of a confocal microscope to observe.

2. Image Acquisition 1. Set up the microscope and computer program. An Olympus confocal microscope equipped with a 100 1.35 NA objective and an Ultraview spinning disc confocal head is controlled by the program Ultraview. The exposure time is set to 300 ms. 2. Find a worm based on the labeled mark and focus on the animal’s head for amphids or the tail for the phasmids. 3. For the time lapse, images are collected for 3 min. And for each worm strain, at least three worms are recorded. As for the Z-stack imaging, the upper and the lower limits of an amphid are set. The interval between each section sets to 0.1 µm.

3. Data Analysis for IFT Assay 1. MetaMorph (Molecular Devices Corporation, Sunnyvale, CA, USA) is used to analyze the recorded images: go to File\Open special\Build stack\Quick (or Ctrl þ Q) to open a stack file. 2. Go to Stack\Kymograph, and set line width to 5 in the Kymograph dialog window. Use the single line function in the tool box and draw a line along the cilium in the

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image. Then click “Create” in the Kymograph dialog window to create a kymograph image (representative kymographs can be seen in Fig. 3). 3. Go to Measure\Region Measurements. In the “Region Measurements” window, select “All Regions” in the “Include” down-drag menu. Click “Open Log” and select “Dynamic Data Exchange (DDE),” and then click OK; a Microsoft Excel file is opened. Draw a line along a kymograph track of a particle. By clicking “Log Data,” the values (Distance and Area) will be transferred into a Microsoft Excel file. 4. Go to Measure\Measure Pixel. In the “Measure Pixel” dialog window, click “Config Log” to select “Image name”, “X”, “Y”. And then double click on the lines in the Kymograph drawn in the “Region Measurements”. The values of X, Y will appear in the opened Excel file (The lines can be seen in Fig. 3). 5. Calculate the distance, time, and velocity. Distance = 0.129*(XLine2 – XLine1) *Distance/Area, where 0.129 is the parameter of the 100 objective lens. At 100, 1 pixel = 0.129 µm. Time = 0.3 (YLine2– YLine1), where 0.3 s is the exposure time. Velocity = Distance/Time.

4. Data Analysis for Cilia Image and Length Measurement 1. MetaMorph (Molecular Devices Corporation) is used to analyze the recorded images: go to File\Open special\Build stack\Quick (or Ctrl þ Q) to open a stack file. 2. Go to Process\Stack Arithmetic\Maximum to make a projection image. 3. Go to Measure\Region Measurements. In the “Region Measurements” window, select “All Regions” in the “Include” down-drag menu. Click “Open Log” and select “Dynamic Data Exchange (DDE)”, and then click OK; a Microsoft Excel file is opened. 4. Go to Measure\Measure Pixel. In the “Measure Pixel” dialog window, click “Config Log” to select “Image name,” “X,” “Y.” And then double click on the lines in the Kymograph drawn in the “Region Measurements.” The values of X, Y will appear in the opened Excel file. 5. Calculate the cilium length. Cilium length = 0.129  SQRT(Xline2 – Xline1)2 þ (Yline2-Yline1)2.

E. Expression and Purification of Kinesin-II and Gliding Motility Assay

1. Baculovirus Expression of Kinesin-II 1. Cloning the subunits of kinesin-II into the baculovirus expression vector: cDNAs encoding the kinesin-II subunits, KLP-11, KLP-20, and KAP-1 are amplified by PCR and inserted into Gateway vector pDONR221 (Invitrogen, Carlsblad, CA, USA), and are subsequently cloned into the destination vector, pDEST8 (Invitrogen). The kap-1 gene contains a 6 His Tag which was incorporated by including the 6 His in the 30 end primer. Recombinant bacmids are generated by transforming the pDEST8 vectors containing klp-11, klp-20, and kap-1 into E. coli MAX Efficiency®

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DH10BacTM cells (Invitrogen). The plasmids in these E. coli cells are modified baculovirus DNA molecules called bacmids. Bacmids are isolated from E. coli DH10Bac and used to transfect Sf9 surface cell cultures to generate baculovirus using Cellfectin® reagent (Invitrogen). After two days of infection the first progeny of viral stocks (P1 viral stocks) are collected. At this stage amplification of the P1 viral stocks is done by following the instructions in the Bac-to-Bac® manual (Invitrogen). The titers of the viral stocks are determined using BacPAKTM Baculovirus rapid titer kit following the manufacturer’s instructions (Clontech Mountain View, CA, USA). The unit for titer of the viral stocks is calculated as number of plaque-forming units per milliliter (pfu/ml). Sf9 cells are cultured using serum-free medium Sf-900TM II (Gibco, Carlsblad, CA, USA) to a concentration of 2106 cells/ml. Mix the three baculovirus stocks by taking equal amount of pfu/ml from each and use this to infect Sf9 cell culture. Culture of Sf9 cells are infected with equal multiplicity of infection [MOI: plaque-forming unit (pfu) of virus per Sf9 cell] from each klp-11, klp-20, and kap-1 viral stocks and are then incubated for three days at 27°C and 130 rpm in a shaking incubator. Harvesting is carried out by centrifugation at 1500 rpm (640 g) at 4°C in a Beckman J6-HC centrifuge JS 4.2 type rotor. The cells are collected and washed once, and the final cell pellet is resuspended in a minimum amount of SFM medium to make a dense slurry and this slurry is used to make small frozen balls by slowly dropping into liquid nitrogenfilled ceramic mortar. The frozen Sf9 cells can be used immediately or stored at –80°C.

2. Purification of Kinesin-II 1. Lysis of Sf9 cells: The frozen Sf9 cells are ground using Micro-Dismembrator S (Sartorius Stedim Biotech, Bohemia, NY, USA) using 20 ml Teflon container and metal balls precooled in liquid nitrogen. The ground powder is suspended in lysis buffer. Lysis of the cells can also be carried out by passing the 5% solution of the cells twice through French press as described in Pan et al. (2006). The subsequent steps of purification are performed at 4°C. 2. The ground powder in the lysis buffer is made into a homogenous mixture by mixing with a glass rod for several minutes. The cell homogenate is then centrifuged at 12,000 rpm (17400 g) for 30 min in a Beckman centrifuge JA-20 type rotor. 3. The supernatant is immediately applied to previously equilibrated Talon® column beads (Clontech Mountain View, CA, USA) and shaken for 1 h with gentle inversions. 4. The Talon® beads and crude extract mixture is then centrifuged at 1500 rpm (640 g) for 5 min in a Beckman J6-HC centrifuge using JS 4.2 type rotor. 5. The supernatant is discarded and the beads are washed twice with lysis buffer. In order to facilitate washing, the mixture is shaken by inversions on a shaker for 10 min. 6. The talon beads slurry is then applied to a disposable 5-ml polypropylene column. 7. The column is washed with lysis buffer containing 10 mM imidazole. The protein is then eluted with lysis buffer containing 150 mM imidazole.

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8. Fractions of 1 ml are collected into Eppendorf tubes and analyzed for protein concentration by Bradford’s assay. 9. The fractions containing proteins are pooled and dialyzed against gel filtration column buffer overnight. 10. The dialysate is passed through a Superose 6 (SuperoseTM 10/300 GL) column. Fractions of 1 ml are collected and they are analyzed by SDS-PAGE for the presence of kinesin-II. 11. The fractions containing the purified kinesin-II protein are collected and concentrated using Amicon Ultra centrifugal filters (Millipore, Billerica, MA, USA) to a concentration of 0.5–1 mg/ml. At the end of the described purification protocol milligram amounts of protein can be obtained with high purity. For illustration SDS-PAGE analysis of samples from different steps of a routine purification, sucrose density gradients, and gel filtration are shown in Fig. 5.

3. Gliding Motility Assay 1. Prepare the flow cell as described in Fig. 6A. Two pieces of the double-sided Scotch tape are attached on two edges of a slide. And then a microscope cover slip with size of 22  22 mm is attached on the tapes to make a flow cell. 2. Use a micropipette to flow in 5 µl of kinesin-II (0.5–1 mg/ml) into the flow cell and incubate for 5 min at room temperature. In the mean time prepare the reaction buffer according to Table III. (The fluorescent microtubules are freshly prepared using Fluorescent Microtubules Biochem Kit following the manufacturer’s instructions (Cytoskeleton, Denver, CO, USA)). Do not allow the flow cell to dry out at any time, allow some extra solution to go out of the flow cell as small droplets. Do not allow air bubbles to be trapped in the flow cell. (Note: When a mixture of heterotrimeric kinesin-II and OSM-3 is used the molarities are used to calculate the ratio of these proteins. The mixture of the proteins should be made before applying to the flow cell by mixing determined moles of two motor proteins.) 3. Flow in 10 µl of blocking solution into the flow cell while touching the opposite side of the flow cell with a piece of filter paper to help flow but not wick off all the solution, and then incubate for 5 min. The set up is shown in Fig. 6B. As the fluid is sucked from one side, the solution pipetted from the other side should fill the flow cell. 4. Flow in 20 µl of reaction buffer and observe under microscope immediately. Seal the flow cell with Valap on both sides. 5. Adjust the focus under microscope (Nikon Eclipse E600 microscope with a digital camera (SenSys from Photometrics, Tucson, AZ, USA) attached and connected to a computer) by aiming the side of the well where you can see the tape edge. Microtubules can be seen when the focus is right. Record time lapse images of the motility for 3 min with 7- to 10-s intervals. 6. Data analysis is performed by using MetaMorph software (Molecular Devices Corporation). This method was used to assay the speed of purified kinesin-II and

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(A)

250 kDa 150 kDa 100 kDa 75 kDa

50 kDa

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97 kDa 66 kDa

(C) 97 kDa 66 kDa

Fig. 5 Preparation and purification of kinesin-II. (A) SDS-PAGE analysis of samples from different steps obtained during the purification of kinesin-II. Lanes: (1) molecular weight marker, (2) high-speed supernatant, (3) supernatant after application to Talon column, (4) rinse with lysis buffer, (5) proteins obtained by washing with 10 mM imidazole containing lysis buffer, (6) Talon column eluate, (7) superose 6 purified kinesin-II. On sucrose density gradients (B) and gel filtration columns (C), the KLP-11, KAP-1, and KLP-20 subunits elute as a monodisperse heterotrimeric complex (S value = 9.8; Rs = 7.1 nm; and native molecular mass = 287 kDa) in a KLP-11/KLP-20/KAP-1 molar stoichiometry of 1.0:1.17:0.89 (protein standard peak positions are also indicated). (Panels B and C are reprinted from Pan et al., 2006.)

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(A)

(B) Cover slip

Glass slide

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Fig. 6 Preparation of the flow cell and the experimental set up for in vitro motility assays. (A) Two pieces of Scotch tape are stuck to the glass slide, leaving a 3-mm gap in between them. Then place a cover slip on top of the tape, gently push down on the cover slip to make it stick, and remove any air bubbles trapped in between the tape, slide and cover slip. Remove the Scotch tape sticking out by peeling off. (B). (1) Flow in the protein solution; the fluid will flow into the cell by capillary action. (2) When adding subsequent solutions, use a filter paper to withdraw the previous solution from one side while replacing it with the next solution by drawing it into the chamber from the other side of the flow cell. The protein sticks to both the cover slip and the glass slide. Focus the microscope on the cover slip in order to see the microtubules being moved by motor protein.

Table III The Reaction Mixture Composition for Gliding Motility Assay Ingredients 2 BRB80 ddH2O 10 Antifade 1 M Phosphocreatine 1000 U/ml Creatine Phosphokinase 2 mM Taxol 50 mM MgCl2 50 mM MgATP Microtubule

Volume (µl) 1.5 12.1 1.0 0.5 0.2 0.2 2.0 2.0 0.5

The volume of the microtubules added can be adjusted depending on the concentration of the MT stock.

OSM-3 individually and also to assay mixtures of OSM-3 and kinesin-II at varying molar ratios ([mol OSM-3]  [mol OSM-3 þ mol kinesin-II]) in competitive motility assays, to determine how these two types of anterograde motor could cooperate to move IFT particles along the cilium (Table II; Fig. 7) (Pan et al., 2006).

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R2=0.99

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0.2 0.4 0.6 0.8 Mole fraction of OSM-3 (WT)

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Fig. 7 Result of competitive MT gliding assays for the two anterograde IFT motors, kinesin-II, and OSM-3. MT gliding rate versus mole fraction (i.e., [mol OSM-3]  [mol OSM-3 þ mol kinesin-II], ignoring solvent concentrations) for mixtures of OSM-3 and kinesin-II. Gliding assay rates were plotted versus mole fraction of wild-type (WT) OSM-3. Experimental data (black dots) with standard deviations (error bars) are shown with best fits for one of two proposed models for the mechanism of coordination of the two motors (curved line). For details see (Pan et al., 2006).

IV. Materials A. Maintenance of WT and dyf Mutant Worms M9 buffer: 6 g Na2HPO4, 3 g KH2PO4, 5 g NaCl, 0.25 g MgSO47H2O, ddH2O to 1 l. Freezing solution: 20 ml of 1 N NaCl, 10 ml of 1 M potassium phosphate buffer (pH 6.0), 60 ml of 100% glycerol, add ddH2O to 200 ml. Autoclave and then add 0.6 ml of 0.1 M MgSO4. B. Genetic Screen of dyf Mutants and Dyf Assay EMS. Fluorescent dye stock solution: 2 mg/ml in DMF (dimethyl formamide). DiI, 1,10 dioctadecyl-3,3,30 ,30 -tetramethylindocarbocyanine perchlorate (Molecular Probes, Eugene, OR, USA), DiO, 3,30 -dioctadecyloxacarbocyanine perchlorate (Molecular Probes). C. Examination of Ciliary Structure Chemicals: anhydrous acetone, 1% osmium tetroxide, 0.1% and 2% uranyl acetate, Epon-Araldite resin, Formvar, Reynold’s lead citrate. Apparatus: high pressure freezer,

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freeze substitution device, 70°C incubator, ultramicrotome with glass or diamond knife, 3mm copper TEM slot grid (Ted Pella Inc., Redding, CA, USA) and a Transmission Electron Microscope. Software: IMOD (freely available from http://bio3d.colorado.edu/imod). D. In Vivo IFT Motility Assay and Cilia Imaging Animals: transgenic worms carrying fluorescently tagged genes. Chemicals: agarose (SeaKem® GTG®, Lonza, Rockland, ME, USA), levamisole [(-)-tetramisole hydrochloride, Sigma, St Louis, MO, USA], Valap (equal weight of lanolin, vaseline, and paraffin). Apparatus: dissecting microscope, confocal microscope with spinning disc. Softwares: Ultraview, MetaMorph (Molecular Devices Corporation). E. Purification of Kinesin-II and Gliding Motility Assay Lysis buffer: 50 mM pipes (pH 6.9), 300 mM NaCl, 1 mM MgCl2, 1 mM b-mercaptoethanol, and EDTA-free protease inhibitor tablet (Roche, Indianapolis, IN, USA). Gel filtration buffer (pH 6.9): 80 mM pipes, 200 mM NaCl, 1 mM MgCl2, 1 mM EGTA, 1 mM DTT, and 0.1 mM ATP. Slides: Gold Seal® Micro slides (Gold Seal Products, Porthsmouth, NH, USA). Cover slips: FisherBrand Microscope cover glass, 2222 mm (Fisher Scientific Pittsburgh, PA, USA). Double-sided scotch tape. Blocking solution: 5 mg/ml casein in 1 BRB80 (Sigma). The stock solutions of the ingredients used in the assay are 2 BRB80: 160 mM pipes (pH 6.9), 2 mM MgCl2, 2 mM EGTA. 10 Antifade (Cytoskeleton): 1 M phosphocreatine, 1 kU/ml creatine phosphokinase, 2 mM Taxol in DMSO (Cytoskeleton), 50 mM MgCl2, 50 mM MgATP, 0.5 µl MT; except 2 BRB80 buffer and MT solution, all the solutions for the gliding motility assay can be prepared in advance, snap frozen in liquid nitrogen, and stored at –80°C in aliquots.

V. Discussion A. Genetic Screen of IFT Mutants EMS mutagenesis, transposon insertion, and TMP-UV deletion provide complementary and valuable approaches for the genetic analysis of IFT and ciliogenesis in C. elegans. EMS mutagenesis is characterized by high, unbiased mutagenicity with low mortality. It was the first method used to generate ciliary mutations in C. elegans and was most widely used procedure to create mutants in the past. Although the subsequent cloning of EMS-induced mutants is labor-intensive, it is still irreplaceable in many cases, since it is the only well-established method for reproducibly creating point mutations in a gene of interest. Some point mutations in the target protein have yielded key structural and functional insights into that protein. For example, osm-3 (sa125) bears a G444/E mutation caused by EMS in the hinge region of the stalk of the OSM-3 protein (Snow et al., 2004), and this mutated protein is constitutively active in in vitro single-molecule motility assays whereas the wild-type OSM-3 protein is autoinhibited and requires proper activation for motility (Imanishi et al., 2006).

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Thus, EMS mutagenesis paved the way to discover a de-repression phenomenon of MT-based motor protein action. Mos1 transposon-mediated insertion mutagenesis has two advantages over EMS mutagenesis: (1) it is biologically safe to humans and (2) it is easy to clone the mutated genes by inverse PCR using Mos1 as a molecular tag (Boulin and Bessereau, 2007). Reverse genetics has been widely used in the postgenomic era, which significantly accelerates the process of studying the functions of candidate ciliary genes. It creates ciliary mutants whose phenotypes are often too subtle to be uncovered by forward genetics. For instance, mutations in KAP-1 and KLP-11 subunits of kinesin-II were firstly isolated by reverse genetics. Although it ultimately turned out that both mutations increase the transport speeds of IFT, and lead to complete loss of axonemes when combined with osm-3 mutants, the mutant animals display superficially normal cilia using the criteria of dye-filling assays and animal behavioral assays, making them difficult to detect (Snow et al., 2004).

B. Examination of Ciliary Structure Serial section TEM is a powerful tool for observing cellular ultrastructure. When coupled with light microscopy, the technique adopts the added capability of correlating dynamic, functional observations with static, yet high-resolution structural analysis. One limitation of this technique is the inability to detect the presence of specific and individual proteins unless they can be labeled by antibody decoration. However, when used to detect changes between wild-type and mutant organisms, these techniques provide a convenient framework for deciphering the functional role of individual proteins and how that function relates to ciliary structure.

C. In Vivo IFT Motility Assay The powerful in vivo IFT assay has dramatically advanced our understanding of the roles and mechanisms of action of IFT motors and IFT particle components and it will continue to do so. However, the limiting factor of this technique is that it does not provide information on whether IFT proteins are moving individually or whether they are moving as components of an assembled complex. Although it is assumed that all the IFT particle components move in a single complex based on the coisolation of the two IFT subcomplexes in C. reinhardtii, it is clear that some IFT components move at a different rate from other IFT components. One example is seen in the bbs mutant background, where IFT-A and IFT-B move separately, with IFT-A moving at the slow rate characteristic of kinesin-II and IFT-B moves at the fast rate characteristic of OSM3 (Ou et al., 2005b). However, it is hard to explain why DYF-2, an ortholog of the IFT144 subunit of IFT-A, moves at the intermediate rate (0.7 µm/s) in the bbs mutant background (Efimenko et al., 2006). Another example that is hard to reconcile with existing models for IFT occurs in the mutant nphp-4, a C. elegans ortholog of nephrocystin, where osm-6::GFP moves at the slow rate (0.5 µm/s) while all other IFT components move at the intermediate rate (0.7 µm/s) (Jauregui et al., 2008). These

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exceptions challenge the models proposed so far and suggest that much more work is needed to better understand the mechanism of IFT in this system. D. In Vitro Gliding Motility Assay The low levels of native motor proteins in C. elegans and lack of feasibility for their purification from this system encouraged IFT researchers to use heterologously expressed proteins. In our experience, the baculovirus expression system has proved to be very useful for producing the complex of three different subunits of heterotrimeric kinesin-II in a high yield and an active form. This system should be useful for expressing and purifying other components of the IFT particles. Biochemical data on the IFT particle complex rely substantially on data from C. reinhardtii (Cole et al., 1998; Piperno and Mead, 1997). Conversely, data obtained from C. elegans by the analysis of rates of IFT in IFT mutants nicely complement biochemical data from C. reinhardtii (Ou et al., 2005b). The in vitro reconstitution of IFT particles can, in principle, be achieved by coexpressing IFT particle components in the baculovirus system for protein purification, which may in turn allow the future reconstitution of the entire process of IFT using in vitro motility assays.

VI. Summary We have summarized the methods that have so far been employed to investigate IFT motors and particles in C. elegans sensory cilia. These methods are included in four subjects: (1) forward and reverse genetics; (2) ciliary structure examination by TEM or fluorescence microscopy; (3) in vivo IFT assays; and (4) heterologous expression of motor proteins for in vitro motility assays. We provide detailed protocols for these methods. We also discuss some of the advantages and disadvantages of these methods which make C. elegans such a powerful system for studying IFT and cilium biogenesis. Acknowledgments Work on IFT research in the Scholey laboratory is supported by NIH grant GM50718. G. Ou is currently supported by a postdoctoral fellowship from the Damon Runyon Cancer Research Foundation in the Vale laboratory at UCSF.

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